G C A T
T A C G
G C A T
genes
Review
RNA Editing and Its Molecular Mechanism in
Plant Organelles
Mizuho Ichinose 1,2 and Mamoru Sugita 1, *
1
2
*
Center for Gene Research, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan;
[email protected]
Institute of Transformative Bio-Molecules, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan
Correspondence: [email protected]; Tel.: +81-52-789-3080
Academic Editor: H. Ulrich Göringer
Received: 24 October 2016; Accepted: 20 December 2016; Published: 23 December 2016
Abstract: RNA editing by cytidine (C) to uridine (U) conversions is widespread in plant mitochondria
and chloroplasts. In some plant taxa, “reverse” U-to-C editing also occurs. However, to date, no
instance of RNA editing has yet been reported in green algae and the complex thalloid liverworts.
RNA editing may have evolved in early land plants 450 million years ago. However, in some plant
species, including the liverwort, Marchantia polymorpha, editing may have been lost during evolution.
Most RNA editing events can restore the evolutionarily conserved amino acid residues in mRNAs or
create translation start and stop codons. Therefore, RNA editing is an essential process to maintain
genetic information at the RNA level. Individual RNA editing sites are recognized by plant-specific
pentatricopeptide repeat (PPR) proteins that are encoded in the nuclear genome. These PPR proteins
are characterized by repeat elements that bind specifically to RNA sequences upstream of target
editing sites. In flowering plants, non-PPR proteins also participate in multiple RNA editing events as
auxiliary factors. C-to-U editing can be explained by cytidine deamination. The proteins discovered to
date are important factors for RNA editing but a bona fide RNA editing enzyme has yet to be identified.
Keywords: RNA editing; chloroplasts; mitochondria; plant organelles; C-to-U editing; U-to-C editing;
pentatricopeptide repeat (PPR) protein; site-recognition specificity factor; cytidine deaminase
1. Introduction
RNA editing is a posttranscriptional modification to nuclear, mitochondrial or chloroplast
genome-encoded transcripts, and occurs in a wide range of organisms. It was discovered in 1986 in
Trypanosoma brucei where uridines were inserted at specific sites in the mitochondrial (kinetoplast)
cytochrome c oxidase II (coxII) transcript to restore the proper protein-coding sequence [1], followed
by a report that described deletion of uridines in coxIII mRNA [2]. This process required guide
RNAs encoded by kinetoplast genomes [3]. Similarly, mitochondrial RNAs in the slime mold
Physarum polycephalum are heavily edited by the insertion of mononucleotides and dinucleotides
at specific sites [4]. In addition, A deletions and nucleotide conversions have also been reported.
Unlike U-insertion/deletion in kinetoplasts, nuclear-encoded transcripts have been shown to undergo
different types of editing; e.g., conversion of cytidine to uridine (C-to-U) in apolipoprotein-B48 mRNA
in human and rabbit intestines [5] and adenosine (A)-to-inosine (I) editing in case of GluR-B mRNA
encoding a glutamate receptor B of glutamate-gated channels [6]. A-to-I editing has also been reported
for several other animal pre-mRNAs. For insights into various types of RNA editing and their
respective mechanistic aspects refer to other review articles in this issue.
In the plant kingdom, RNA editing was first identified as a C-to-U exchange in mitochondrial
transcripts in 1989 [7–9], followed by its reporting in chloroplasts, two years later [10]. RNA editing
occurs mostly in translated regions of organelle mRNAs, and occasionally, also in the untranslated
Genes 2017, 8, 5; doi:10.3390/genes8010005
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Genes 2017, 8, 5
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regions, introns and structural RNAs [11,12]. Most of the C-to-U changes in the protein-coding region
lead to preservation of evolutionarily codons. In some plant taxa, U-to-C “reverse” editing has also
been frequently reported in both plant organelles. Therefore, RNA editing is believed to serve as
a correction mechanism at the post-transcriptional level for T-to-C (or C-to-T) mutations, probably
acting as buffer to less favored mutations in the genomic coding sequences [11,12]. Across plant
kingdom, the frequency of organellar RNA editing varies from zero to thousands of sites. No editing
seems to occur in nuclear genome-encoded transcripts in plants. Comprehensive and excellent review
articles on plant RNA editing have recently been published and describing its mechanistic and
functional aspects [13–17]. Here, we briefly summarize the RNA editing events in green plant lineages
and current knowledge of trans-acting factors involved in C-to-U RNA editing in chloroplasts and
plant mitochondria.
2. RNA Editing Events in Plant Organelles
2.1. C-to-U RNA Editing
RNA editing sites in translated regions can be predicted by a comparison of amino acid
sequences deduced from genomic DNA sequences from various plant species. Subsequently, RNA
editing can be verified by cDNA sequence analysis. A number of editing sites identified in various
land plant mitochondria and chloroplasts [18–43] are listed in Table 1. There are 20 to 60 editing
sites in chloroplasts and 300 to 600 sites in mitochondria of most flowering plants, except for the
early-branching flowering plant Amborella trichopoda. In seed plants, all these editing events are of
C-to-U type. Most of the sites in translated regions are efficiently edited, with 90%–100% efficiency,
in green leaves. On the other hand, the efficiency of C-to-U editing events that create a translation
initiation codon (by an ACG to AUG change) has been surprisingly low. For instance, the editing
efficiency at the ndhD-1 site in the Arabidopsis chloroplast ndhD transcript is 45% [21] and that of
the rps14-C2 site in the moss Physcomitrella patens chloroplast rps14 mRNA is 70% in filamentous
protonemata, which reduces further to only 20% in leafy tissues [30]. This suggests that editing at this
site may regulate translation in chloroplasts. RNA editing efficiency varies in different tissues and
organs, developmental stages, or different mutant lines [44].
Table 1. The numbers of RNA editing sites in chloroplasts and plant mitochondria.
Plant Species (Common Name)
Chloroplasts
Seed plants (monocotyledonous angiosperms)
Oryza sativum (rice)
Zea mays (maize)
Spirodela polyrhiza (greater duckweed)
Seed plants (dicotyledonous angiosperms)
Arabidopsis thaliana (thale cress)
Nicotiana tabacum (tobacco)
Cucumis sativus (cucumber)
Amborella trichopoda
Seed plant (gymnosperms)
Cycas taitungensis (Emperor Sago)
Ferns
Adiantum capillus-veneris (southern maidenhair fern)
Ophioglossum californicum (California adder’s tongue fern)
Psilotum nudum (whisk fern)
Equisetum hyemale (horsetail)
Lycophytes
Selaginella uncinata (spike moss)
Bryophytes
Anthoceros angustus (hornwort)
Physcomitrella patens (moss)
Marchantia polymorpha (liverwort)
RNA Editing Type
C-to-U
U-to-C
References
21
26
66 *
0
0
0
[18]
[19]
[20]
43 *
34
51
138
0
0
0
0
[21]
[22]
[23]
[23]
85
0
[24]
315
297
27
0
35
3
0
0
[25]
[26]
[26]
[27]
3415 *
0
[28]
509
2
0
433
0
0
[29]
[30]
Genes 2017, 8, 5
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Table 1. Cont.
Plant Species (Common Name)
Mitochondria
Seed plants (monocotyledonous angiosperms)
Oryza sativum
Seed plants (dicotyledonous angiosperms)
Arabidopsis thaliana
Brassica napus L. (rapeseed)
Beta vulgaris (sugarbeet)
Vitis vinifera (grapevine)
Phoenix dactylifera L. (date palm)
Nicotiana tabacum
Seed plant (gymnosperms)
Cycas taitungensis
Lycophytes
Isoetes engelmannii (Engelmann0 s quillwort)
Selaginella moellendorffii (spike moss)
Bryophytes
Physcomitrella patens
Marchantia polymorpha
RNA Editing Type
C-to-U
U-to-C
References
491
0
[31]
619 *
427
357
445 *
592
635 *
0
0
0
0
0
0
[32,33]
[34]
[35]
[36]
[37]
[38]
565
0
[39]
1560 *
2152 *
222 *
0
[40]
[41]
11
0
0
0
[42,43]
Numbers of editing sites in species in which full complement have been analysed. * Data from RNA-seq analyses.
Recent high-throughput RNA-seq analyses have revealed minor RNA editing events in
untranslated regions and intron sequences as well as in protein-coding regions. For instance, in
addition to the 34 already known editing sites in Arabidopsis chloroplasts [45], nine novel sites have
been identified that are edited at a low level (5% to 12%) [21]. Among the 635 identified editing sites in
Nicotiana tabacum mitochondria, five sites are in tRNAs and 73 in non-coding regions [38]. Across the
plant kingdom, the total number of C-to-U editing sites in chloroplasts varies from 0 in the liverwort
Marchantia polymorpha to 3415 in the spike moss Selaginella uncinata [28] (Table 1).
Out of 3415 sites identified in 74 S. uncinata chloroplast mRNAs, 428 are silent editing events,
74 have been identified in four group II introns, 52 create start codons and 31 create stop codons [28].
A total of 2139 editing sites in 18 mRNAs were identified in S. moellendorffii mitochondria [41]. Of these,
424 are silent, whereas the others result in 1488 codon changes. In addition, 13 sites are in the two
rRNAs [41]. To date, RNA editing sites can be predicted by Plant RNA-editing prediction and analysis
computer tools PREPACT 2.0 [46] and PREP-Mt [47]. Some 1800 C-to-U editing sites have been
predicted in the S. moellendorffii chloroplast, 460 sites in the quillwort Isoetes flaccida (chloroplast) and
340 sites in Huperzia lucidula chloroplasts [28]. Therefore, the organellar transcripts in Selaginella, one
of the early vascular plant lycopods, seem to be most commonly edited.
In case of the bryophyte (early non-vascular land plants) P. patens, where there are only two
identified C-to-U editing sites in chloroplasts [30], there are 11 such site in its mitochondria [42,43]. On
the other hand, hornworts such as Anthoceros and Phaeoceros laevis undergo substantial RNA editing [29,
48]. However, no editing event has so far been reported in green algae, including Chara vulgaris
(stonewort), suggesting that the process of RNA editing may have evolved only after the plants
established themselves on the land.
2.2. U-to-C RNA Editing
Although in none of the seed plants listed in Table 1, U-to-C RNA editing has been reported in
either of the two organelles, a 25 year old report describes U-to-C editing in plant mitochondria in
wheat cox3, evening primrose cob and cox2 and pea cox2 transcripts (references in [11]). While these
data need to be reanalyzed, the rare instances of reverse-type editing occurring in seed plants cannot
to completely ruled out. In contrast to higher plants, the reverse (U-to-C) editing appears to be
restricted to hornworts, lycophytes, and ferns [27] with an extensive array of (over 400) U-to-C editing
sites identified in the hornwort, Anthoceros angustus, chloroplasts [29] and mitochondria of two other
Genes 2017, 8, 5
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hornwort species [48] as well as in the early vascular plant lycophyte Isoetes engelmannii [40]. The cDNA
sequence analysis of four selected mitochondrial genes from nearly 30 ferns species has revealed that
both types of editing is prevalent in most fern species, and notably, the reverse (U-to-C) editing could
even exceed the C-to-U RNA editing in some ferns [27]. For instance, 53 C-to-U and 70 U-to-C editing
sites were detected in the 1020 bp cDNA sequence of the mitochondrial atp1 gene of Anemia phyllitidis.
In other ferns such as Equisetum hyemale, several C-to-U editing sites but no U-to-C editing have
been found in the selected gene transcripts. In E. hyemale chloroplasts, the RNA editing is completely
absent [27]. It would be interesting to know the reasons for the expansion of U-to-C type of editing
only in some specific plant taxa, including the Monilophytes. The evolution of editing in Monilophytes
spans a much longer timeframe, probably as old as the seed plants. Hornworts, some lycopytes, and
ferns produce spores but not seeds. It is possible that seed plants and some bryophytes might have
lost the U-to-C editing during the course of evolution.
3. RNA Editing Affects tRNA Maturation and RNA Splicing
RNA editing in plant organelles mostly affects mRNAs, thus providing the means to correct
genetic information for proper protein function. In addition, editing affects some tRNAs and rRNAs
encoded in the organellar genomes [49–52]. In bean and potato mitochondria, a C-to-U editing event
corrects a C:A mismatch base pair into a U:A base pair in the acceptor stem of tRNAPhe [51]. In larch,
three C-to-U editing events restore U:A base pairs in the acceptor, D and anticodon stem, respectively,
in mitochondrial tRNAHis [52]. In the lycophyte I. engelmanni mitochondria, ten tRNAs are edited
to improve base pairing in stem regions [40]. Thus, editing events in pre-tRNAs help in restoring
the RNA secondary structure by removing mismatches in the double-stranded stem region and are
a prerequisite for their processing into functional tRNAs.
Exemplifying the rarity of editing in rRNAs in seed plants, there was no such report for a long
time after an initial reporting of two potential sites in Oenothera mitochondrial 26S rRNA 25 years
ago [50]. Recently however, 13 C-to-U editing sites have been identified in two rRNAs in the lycophyte
S. moellendorffii mitochondria [41]. Three of these sites are in the 26S rRNA, and rest 10 are in the first
exon of 18S rRNA. Notably, RNA editing at the last nucleotide of the 18S rRNA 50 exon may directly
influence splicing of its group I intron, as it likely forms the U:A base pairing needed for the conserved
paired region P1 [41].
Like some instances in tRNAs, perhaps the editing sites within group II introns are also of
functional importance because editing can improve the base pairing required for splicing. Domain VI
of nad1 group II intron 3 from Oenothera mitochondria is modified by C-to-U editing to generate
the typical domain VI secondary structure. Self-splicing in vitro is observed only in the edited (A:U
basepair) form, indicating that this editing event is a prerequisite for splicing [53]. In the lycophyte S.
uncinata chloroplasts, a number of intron editing events have been identified, which could possibly
improve the RNA secondary structure of group II introns, including the highly conserved domains
V of the intron 30 termini [28]. Such editing events could potentially play significant role in splicing,
thereby regulating the availability of functional tRNAs.
RNA editing in exons close to splice sites may also affect intron splicing or vice versa. For instance,
the spinach chloroplast ndhA mRNA is edited at two sites, one of which is located only 12 nucleotides
downstream of the 30 intron-exon splice site. To assess if RNA editing occured after or before splicing,
short “spliced” and “unspliced” ndhA gene fragments were introduced and transcribed within tobacco
chloroplasts. The subsequent cDNA analysis showed that only spliced ndhA mRNAs were edited [54].
A similar result was observed in case of the moss P. patens mitochondrial atp9 gene [55]. This atp9 gene
is interrupted by three introns and an editing site lies within the third exon (only 8 nt long). This site
is completely edited in fully spliced mRNA, while it remains unedited in the unspliced mRNA [55].
These observations suggest that splicing precedes editing. In contrast, the land plant chloroplast
tRNALeu gene contains a group I intron between the first and second position of the UAA anticodon.
In the moss Takakia lepidozioides, the CAA anticodon of tRNALeu is edited to create a canonical UAA
Genes 2017, 8, 5
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codon [56]. The spliced tRNALeu is completely edited while unspliced tRNAs are partially edited.
This suggests that the anticodon editing of tRNALeu may occur before RNA splicing.
4. RNA Editing Factors in Plant Organelles
4.1. PPR Proteins as a Site-Recognition Factor
To elucidate the molecular mechanism of RNA editing in plant organelles, in vitro, in vivo, and
in organello studies have extensively been performed using flowering plants such as wheat, tobacco,
pea, and cauliflower [57–61]. These studies have helped in identifying cis-acting elements adjacent to
editing sites and discovering putative site-specific proteins that interact with these elements. In all
such instances, the cis-elements comprise stretches of 20 to 25 nucleotides upstream of the editing
sites. The identification of the first trans-acting factor, however, was not the outcome of any genetic
and biochemical study for editing factors, but instead it resulted from a study on photosynthetic
mutants in 2005 [62]. One of the isolated mutants, chlororespiratory reduction 4 (crr4), showed defects in
the accumulation of the plastidic NADH dehydrogenase (NDH) complex, which is a multi-subunit
complex in the thylakoid membrane. The loss of NDH complex was correlated directly to the loss of
a C-to-U editing event that otherwise creates the start codon AUG in ndhD mRNA. It was later found
that the CRR4, a member of the pentatricopeptide repeat (PPR) protein family, binds to a 36 nucleotides
(−25 to +10) region surrounding its target editing site [63]. This suggested that CRR4 could be the
bona fide trans-acting factor essential for recognizing this RNA editing target site. Following this
discovery, several other PPR proteins were identified as site recognition factors affecting editing
in chloroplasts and mitochondria [64,65] (Figure 1). Many editing PPR proteins were found to be
responsible for only a single editing site, whereas, some PPR proteins could recognize multiple sites
with similar cis-element sequences [13–17].
PPR proteins constitute a large family of nuclear-encoded proteins comprising of 100 to over
1000 members in land plants [66–68]. However, there number varies from only several to 20 members
in fungi, protists, and animals [66]. Almost all the PPR proteins are localized in either chloroplasts or
mitochondria, or both [69] where these proteins participate in different facets of RNA metabolism such
as RNA splicing, RNA stability, and translational initiation [70]. PPR proteins are characterized by
tandem arrays of the degenerate 31 to 36-amino acid PPR motif that folds into a pair of anti-parallel
Genes 2017, 8, 5 6 of 15 alpha helices, which have been suggested to specifically bind to RNA sequence targets [71].
Figure 1.
pentatricopeptide
repeat
(PPR)
editing
proteins
and a and model
theirfor binding
Figure 1. Plant
Plant organellar
organellar pentatricopeptide repeat (PPR) editing proteins a for
model their to the editing site. Schematic domain structure of PPR editing proteins that consist of PPR motifs
binding to the editing site. Schematic domain structure of PPR editing proteins that consist of PPR (P,
L, S),(P, and
C-terminal
domains domains (E and DYW).
The
DYWThe domain
contains
the
conserved
motifs L, additional
S), and additional C‐terminal (E and DYW). DYW domain contains the n CxxC. The PPR tract interacts with a target RNA in a one PPR
zinc-binding
motif
signature,
HxE(x)
n
conserved zinc‐binding motif signature, HxE(x) CxxC. The PPR tract interacts with a target RNA in motif to one nucleotide manner. The last PPR S motif recognizes nucleotide at position –4 from the
a one PPR motif to one nucleotide manner. The last PPR S motif recognizes nucleotide at position –4 from editing site (+1).
the editing site (+1). Recently, a code are
for structurally
PPR‐RNA divided
recognition has major
been classes,
elucidated [75–77]. acid The PPR proteins
into two
denoted
P andThe PLS.amino The P-class
combinatorial patterns at position 6 and 1ʹ (position 1 of the following PPR motif) recognize a specific is composed of canonical PPR (P) motifs of 35 amino acids, while the PLS-class consists of canonical P
RNA base (Figure 2). The PPR editing proteins bind to a specific cis‐element for editing. PPR crystal motifs and their variants L (for long, 35 or 36 amino acids) and S (for short, 31 amino acids), which
structure analyses have shown that a PPR in N‐to‐C terminus orientation interacts with RNA in the 5′ to 3′ orientation for the target RNA and confirms the RNA‐binding code [78–81]. Similarly, PPR editing proteins bind to specific cis‐elements for editing in a one‐PPR motif to one‐nucleotide manner. Genes 2017, 8, 5
6 of 15
differ in sequence length and conservation [66,70]. At their C-terminus, following the last PPR motif,
many PLS-class PPR proteins are extended by the plant-specific conserved E (extension)
domain, and
are thus occasionally called PPR-E or E-type PPR proteins. The Arabidopsis CRR4 also belongs to this
category.
About
halforganellar of the PLS-class
PPR proteins
with
the Eediting domain
are further
a DYW
Figure 1. Plant pentatricopeptide repeat (PPR) proteins and a extended
model for by
their binding to the editing site. Schematic domain structure of PPR editing proteins that consist of PPR domain
of about 100 amino acids and are named after its three highly conserved C-terminal amino
(P, L, S), (D),
and tyrosine
additional C‐terminal domains (W).
(E and DYW). The DYW the only
acids,motifs aspartic
acid
(Y),
and tryptophan
The
PLS-class
PPR domain proteinscontains are found
nCxxC. The PPR tract interacts with a target RNA in conserved zinc‐binding motif signature, HxE(x)
in land
plant lineages but not in algae and non-plants.
Intriguingly, DYW-type PPR proteins were
founda one PPR motif to one nucleotide manner. The last PPR S motif recognizes nucleotide at position –4 from in the protist microscopic amoeba Naegleria, in the slime mold Physarum and in the wheel animal
the editing site (+1). Rotifera
[72–74]. To date, nearly 70 PPR editing factors have been identified in seed plants and the moss,
P. patens [15,16]. All of them belong to the PLS-class with C-terminal E or E-DYW domains (Figure 1).
Recently, aa code
code for
for PPR-RNA
PPR‐RNA recognition
recognition has
has been
been elucidated
elucidated [75–77].
[75–77]. The amino acid
acid Recently,
The amino
combinatorial patterns at position 6 and 1ʹ (position 1 of the following PPR motif) recognize a specific 0
combinatorial patterns at position 6 and 1 (position 1 of the following PPR motif) recognize a specific
RNA base (Figure 2). The PPR editing proteins bind to a specific cis‐element for editing. PPR crystal RNA
base (Figure 2). The PPR editing proteins bind to a specific cis-element for editing. PPR crystal
structure analyses have shown that a PPR in N‐to‐C terminus orientation interacts with RNA in the structure analyses have shown that a PPR in N-to-C terminus orientation interacts with RNA in the
0 to 30 orientation for the target RNA and confirms the RNA-binding code [78–81]. Similarly, PPR
55′ to 3′ orientation for the target RNA and confirms the RNA‐binding code [78–81]. Similarly, PPR editing proteins bind to specific cis‐elements for editing in a one‐PPR motif to one‐nucleotide manner. editing
proteins bind to specific cis-elements for editing in a one-PPR motif to one-nucleotide manner.
Figure
2. PPR recognition code for RNA binding. Key amino acid positions 6 and 10 of each PPR motif
Figure 2. PPR recognition code for RNA binding. Key amino acid positions 6 and 1′ of each PPR motif are
indicated as yellow and blue colored square boxes, respectively. T, N, D, and S denote amino acids
are indicated as yellow and blue colored square boxes, respectively. T, N, D, and S denote amino acids tyrosine,
asparagine, aspartic acid, and serine, respectively. Combinations of amino acids at positions 6
tyrosine, asparagine, aspartic acid, and serine, respectively. Combinations of amino acids at positions 0 specify binding to specific bases as proposed in Barkan et al. [75]. (T, N) (T at 6, N at 10 ) specify
and
1
6 and 1′ specify binding to specific bases as proposed in Barkan et al. [75]. (T, N) (T at 6, N at 1′) specify binding
to adenine (A), (T, D) to guanine (G), (N, S) to cytidine (C), (N, D) to uridine (U), and (N, N)
binding to adenine (A), (T, D) to guanine (G), (N, S) to cytidine (C), (N, D) to uridine (U), and (N, N) to
C or U.
to C or U. 4.2.
Importance of the DYW Domain in RNA Editing
4.2. Importance of the DYW Domain in RNA Editing Plant-specific
E domains, which contain two PPR-like motifs, have been shown to be essential for
Plant‐specific E domains, which contain two PPR‐like motifs, have been shown to be essential editing
[64,82].
Okuda
et al.et [83]
that the
DYW
of CRR28
OTP85
for editing [64,82]. Okuda al. demonstrated
[83] demonstrated that the domains
DYW domains of and
CRR28 and interact
OTP85 with
the target C, whereas the E domain of CRR21 is not involved in binding. The exact role of the E
interact with the target C, whereas the E domain of CRR21 is not involved in binding. The exact role domain
editing remains
is speculated
however that
it might
beit involved
ininvolved interacting
of the E in
domain in editing unclear,
remains itunclear, it is speculated however that might be in with
other proteins.
interacting with other proteins. Like
E-type PPR proteins, in most cases a single PPR-DYW editing protein is involved in RNA
Like E‐type PPR proteins, in most cases a single PPR‐DYW editing protein is involved in RNA editing
at
The DYW DYW domains
editing at target
target sites
sites (Figure
(Figure 3a).
3a). The domains contain
contain the
the canonical
canonical zinc-binding
zinc‐binding motif
motif HxE(x)nCxxC,
which is also found in other cytidine deaminases [84]. In addition, there also exists
HxE(x)nCxxC, which is also found in other cytidine deaminases [84]. In addition, there also exists a acorrelation correlation between between the
distribution
of nuclear
DYW DYW domains
and instances
of organelle
the evolutionary
evolutionary distribution of nuclear domains and instances of RNA editing among land plants [85]. Put together, these findings suggest that DYW domain could
harbor the cytidine deaminase enzymatic activity. However, the RNA deamination activity of the DYW
domain has not yet been proven by any of the studies deploying in vitro editing assay systems [82,86].
Furthermore, genetic analyses carried out on CRR22 and CRR28 suggest that the DYW motif is
Genes 2017, 8, 5 7 of 15 organelle RNA editing among land plants [85]. Put together, these findings suggest that DYW domain could harbor the cytidine deaminase enzymatic activity. However, the RNA deamination Genes
2017, 8, 5
7 of 15
activity of the DYW domain has not yet been proven by any of the studies deploying in vitro editing assay systems [82,86]. Furthermore, genetic analyses carried out on CRR22 and CRR28 suggest that the DYW motif is dispensable for the editing activity [82], further accentuating the ambiguity over dispensable for the editing activity [82], further accentuating the ambiguity over the role of the
the role of the DYW domain. DYW
domain.
In contrast to the findings in Physcomitrella, the DYW1 protein in Arabidopsis has been identified In contrast to the findings in Physcomitrella, the DYW1 protein in Arabidopsis has been identified
as an RNA editing factor acting specifically on the chloroplast ndhD‐1 site [87]. DYW1 consists of a as
an RNA editing factor acting specifically on the chloroplast ndhD-1 site [87]. DYW1 consists of
apartial E domain and a well‐conserved DYW domain with no PPR motifs. In chloroplasts, the DYW1 partial E domain and a well-conserved DYW domain with no PPR motifs. In chloroplasts, the DYW1
interacts in with an E‐type PPR protein CRR4 to edit the ACG codon to an AUG in the ndhD mRNA. interacts in with an E-type PPR protein CRR4 to edit the ACG codon to an AUG in the ndhD mRNA.
Both proteins have been shown to be required for this editing event, suggesting that the DYW domain Both
proteins have been shown to be required for this editing event, suggesting that the DYW domain
is essential but could be provided in trans if not present in cis on the PPR editing factor (Figure 3b). is essential but could be provided in trans if not present in cis on the PPR editing factor (Figure 3b).
There are
are five
five other DYW1‐like proteins in Arabidopsis, suggesting that association with DYW There
other
DYW1-like
proteins
in Arabidopsis,
suggesting
that association
with DYW
proteins
proteins could be a general feature of E‐type PPR editing factors [87]. could be a general feature of E-type PPR editing factors [87].
(a)
CRR28, OTP85, PpPPR_56
E
C
U
RNA
(b)
DYW
CRR4
DYW1
E
C
U
RNA
(c)
VAC1
RARE1
E
RNA
DYW
C
U
Figure
3. RNA editing requires a single or multiple PPR editing factors. (a) Single PPR-DYW editing
Figure 3. RNA editing requires a single or multiple PPR editing factors. (a) Single PPR‐DYW editing proteins
(e.g., CRR28, OTP85, PpPPR_56) are involved in editing at their target sites; (b) PPR-E editing
proteins (e.g., CRR28, OTP85, PpPPR_56) are involved in editing at their target sites; (b) PPR‐E editing factor
(e.g.,
CRR4) and DYW1 are both required for editing at a single site. PPR proteins recognize
factor (e.g., CRR4) and DYW1 are both required for editing at a single site. PPR proteins recognize the the
target
editing
and
DYW1
are
involved
transin inediting; editing;(c) (c) Two Two PPR‐DYW PPR-DYW proteins
target editing site site
and DYW1 are involved in intrans proteins are
are cooperatively
involved in editing. Either of two PPR-DYW proteins is involved in site recognition and
cooperatively involved in editing. Either of two PPR‐DYW proteins is involved in site recognition and another
one may be required for the C-to-U editing reaction.
another one may be required for the C‐to‐U editing reaction. The PPR proteins have also been shown to act cooperatively, as the loss‐of‐function of one may The
PPR proteins have also been shown to act cooperatively, as the loss-of-function of one may
reduce but not completely abolish editing at a particular site, suggesting that the remaining editing reduce but not completely abolish editing at a particular site, suggesting that the remaining editing
could be carried out by its other counterpart(s). For instance, RARE1 and VAC1 (also called AtECB2), could
be carried out by its other counterpart(s). For instance, RARE1 and VAC1 (also called AtECB2),
both of which are DYW‐type PPR proteins, are identified as editing PPR proteins targeting the same both
of which are DYW-type PPR proteins, are identified as editing PPR proteins targeting the same
accD‐C794 site in Arabidopsis chloroplasts [88,89]. Mutation of the RARE1 gene results in a complete accD-C794
site in Arabidopsis chloroplasts [88,89]. Mutation of the RARE1 gene results in a complete
loss of accD editing [88] while that of VAC1 leads to a 60% reduction of editing compared to the wild loss
of accD editing [88] while that of VAC1 leads to a 60% reduction of editing compared to the wild
type level [89]. An in silico target assignment test suggested that RARE1, but not VAC1, is indeed a type
level [89]. An in silico target assignment test suggested that RARE1, but not VAC1, is indeed
site‐recognition not be
be a site-recognitionfactor factorfor foraccD accDediting editing[90]. [90].VAC1 VAC1is isinvolved involvedin in accD accD editing, editing, but but might might not
required for site recognition. These two PPR‐DYW proteins could be cooperatively involved in accD required
for site recognition. These two PPR-DYW proteins could be cooperatively involved in accD
editing, and VAC1 may interact with RARE1, as DYW1 does with CRR4 (Figure 3c). editing, and VAC1 may interact with RARE1, as DYW1 does with CRR4 (Figure 3c).
Similarly, studies on moss PPR editing factors also support the importance of the DYW domain Similarly,
studies on moss PPR editing factors also support the importance of the DYW domain in
in RNA editing. The moss (P. patens) genome encodes 10 DYW‐type PPR proteins but no E‐type PPR RNA
editing. The moss (P. patens) genome encodes 10 DYW-type PPR proteins but no E-type PPR protein.
In this moss, the 13 C-to-U editing events are coordinated by nine DYW-type PPR proteins [55,91,92],
implying that one or more PPR-DYW proteins would have to act as a site-recognition factor for more
Genes 2017, 8, 5
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than one editing sites (Figure 3a). This is the first full assignment of nuclear-encoded DYW-type editing
protein factors to all its organellar editing sites in a plant species.
Recently, DYW domains have been shown to bind zinc ions and have been implicated in RNA
editing in Arabidopsis [93,94]. Moreover, in vitro RNA binding assays have shown that DYW domains
in chloroplast editing factors, CRR28 and OTP85, directly bind to their target Cs and respective 50
proximal region from –3 to 0 (+1C) [83]. This suggests their involvement in the C-to-U catalytic
reaction. On the same lines, we have also observed that the zinc-binding motif of DYW-type protein
PpPPR_56 is essential for editing at two mitochondrial sites nad3-C230 and nad4-C272 in P. patens
(Ichinose and Sugita, unpublished). Various transgenes for wild type PpPPR_56 (56comp), and mutant
variants with the HxE(x)nCxxC motif changed to alanine, AxA(x)nCxxC (56M1) and HxE(x)nAxxA
(56M2), respectively, were introduced into the PpPPR_56 knockout moss (∆56-22). In the 56comp moss
line, editing of nad3-C230 and nad4-C272 sites was restored to wild type levels. Whereas, 56M1 and
56M2 constructs failed to complement the mutant editing phenotype, thereby suggesting that the
zinc-binding motif of the DYW domain could also play an important role in the process of editing in
the moss system as well.
4.3. Non-PPR Editing Factors in Plant Organelles
Besides RNA sequence-specific PPR editing factors, another group of proteins, known as multiple
organellar RNA-editing factors (MORFs) have also been linked to RNA editing in flowering plants [95].
MORFs are also known as RNA editing Interacting Proteins (RIPs) [96]. Ten members of the MORF
family, with a novel conserved protein domain, named the MORF domain, have been identified in
Arabidopsis. Seven of these target sites in mitochondria, two (MORF2 and 9) in chloroplasts and one
(MORF8) acts on its targets in both the organelles. In contrast to PPR editing factors, mutants of either
MORF2 or MORF9 gene are affected at almost all RNA editing sites in Arabidopsis chloroplasts [95].
This suggests that editing of the ndhD-1 site requires at least four proteins: CRR4, DYW1, MORF2
and MORF9 [80]. Similarly, mitochondria-localized MORFs are also involved in RNA editing at
many sites. MORF proteins have been shown to interact with each other and also with some PPR
editing factors [97,98] and form specific homo- and heteromeric interactions [99]. These factors are
organized in a higher ordered editing complex (~200 kDa, called the editosome) [96]. Although the
actual function of MORF proteins in the editosome in organelles is as yet unknown, the members of
this family may act as connectors between the PPR editing factors and the actual cytidine deaminase
activity site in the editosome. This hypothesis, however, needs to be validated. The mitochondrial
MORF proteins discriminate between different PPR proteins in yeast two-hybrid assays [95]. In some
instances, the MORF proteins that are required for editing at a given site indeed interact with the
specific PPR protein, which is also essential for processing that particular site. The MORF proteins
may be involved in bridging the distance of four nucleotides between the nucleotides contacted by the
PPR proteins and the actually edited C moiety to guide the enzyme.
Other types of proteins involved in RNA editing belong to RNA recognition motif
(RRM)-containing proteins: chloroplast ribonucleoproteins (cpRNPs) and organelle RRM proteins
(ORRMs). CP31A, a member of the cpRNP family containing two RRMs, influences the efficiency of
editing at 13 sites in Arabidopsis chloroplasts [100]. However, this effect on editing is possibly indirect
because the levels of many other transcripts are also reduced in the cp31a mutant [101]. The ORRM
family proteins have also been shown to be involved in the process of editing in Arabidopsis and
maize [102–104]. ORRM1 is a chloroplast localized protein that is characterized by two truncated
MORF domains and one RRM domain [102]. The loss of ORRM1 leads to a drastic reduction of editing
at 12 sites in Arabidopsis and nine sites in maize chloroplasts. ORRM2, ORRM3 and ORRM4, which
have a single RRM domain and a glycine-rich domain, are likewise shown to be important for efficient
editing at many mitochondrial sites [103,104]. ORRMs have also been shown to associate with MORF
proteins and form homo or heteromeric interactions. In addition, ORRM1 can interact with PPR editing
factors. Similar to MORFs, it seems likely that the ORRMs also are major components of the editosome.
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Two additional novel proteins, PPO1 (protoporphrinogen IX oxidase 1) and OZ (organelle
zinc finger), have been characterized as general editing factors [105,106]. Notably, PPO1, a critical
enzyme for the tetrapyrrole biosynthetic pathway, plays an unexpected role in chloroplast editing
at multiple sites in Arabidopsis [105]. PPO1 interacts with three chloroplast MORF proteins but not
with PPR proteins. These data suggest that PPO1 controls the level of chloroplast editing via the
stabilization of MORFs. The OZ family contains 4 members, OZ1-4, in Arabidopsis of which three
are predicted to be localized in chloroplasts while one is mitochondrial. The OZ1 was identified
by co-immunoprecipitation with DYW-type PPR editing factor, RARE1 [106]. Disruption of OZ1 in
Arabidopsis leads to an alteration in the level of editing of most sites in chloroplasts. OZ1 can interact
with PPR editing factors, where it is assigned to the cognate sites, and ORRM1, but not with MORFs.
This interaction supports the notion that OZ1 takes part in the editosome of chloroplasts.
5. Mechanism of RNA Editing in Plant Organelles
In plant organelles, PLS-type PPR proteins recognize and bind specifically to editing sites.
The PPR–RNA complex is organized into the editosome with several additional non-PPR protein factors
such as MORF and ORRM proteins. However, the order of addition/assembly of individual protein
factors into the editosome has yet to be clarified. These non-PPR proteins might play a key role in editing
as regulators of editing efficiency or as connectors of site-specific PPR proteins with other proteins
or an unidentified editing enzyme. Presumably, approximately 200 PLS-type PPR proteins found in
Arabidopsis might be involved in RNA editing. Henceforth, several of the editing PPR protein would
have to recognize more than one editing sites to be able to recognize all 600 editing events in Arabidopsis.
PPR proteins and non-PPR editing factors are targeted to either chloroplasts or mitochondria, or
both. This suggests that the basic machinery for a C-to-U editing event is perhaps conserved in both
organelles. This editosome model has been drawn from molecular evidence found mostly in Arabidopsis.
However, in the early land plants (mosses) and early vascular plants (lycophytes), the non-PPR editing
factors described above are not encoded in their nuclear genomes. Unlike the complex editosome of
seed plants, RNA editing may occur in a simpler editing complex, composed of a single PPR-DYW
editing protein and a few other unidentified non-PPR editing factors, at least in mosses. The remaining
central question is the nature of the RNA editing enzyme. Despite circumstantial evidence supporting
the DYW editing enzyme [85,86], a biochemical demonstration of cytidine deaminase activity for the
DYW domain would be required to prove that DYW indeed is the editing enzyme.
6. Conclusions and Perspectives
In some plant species, the existence of both conventional C-to-U and reverse U-to-C editing events
is highly evident, but the mechanism of target recognition and features of the editing factors involved
are completely unknown. In the fern A. phyllitidis, there are 53 C-to-U and 70 U-to-C editing events
known in the 1 kb mitochondrial atp1 mRNA [27]. Such a high density, of more than 100 target sites in
a 1 kb transcript, suggests that there would exist several overlapping cis-elements which are needed
to be properly identified in their unedited, partially-edited or fully-edited states. Although C-to-U
editing requires PLS type PPR proteins, it is unclear if the same were true also for the reverse (U-to-C)
editing. However, the possibility of PLS-type PPR proteins being involved in reverse editing cannot
also be ruled out either. About 6000 C-to-U (but no U-to-C) editing sites are present in some Selaginella
species, suggesting that thousands of PPR editing factors could be involved in all editing events.
To determine the cis-elements for editing, an in vitro or in organello assay system must be
developed from the plants in which reverse editing also occurs. However, so far it has been difficult
to prepare purified organelles from these plants. To identify the editing factors, a forward and/or
reverse genetic approach could be useful, as it has been in case of flowering plants. However, such
approaches have yet to be applied. The development of breakthrough technologies for RNA editing
studies in reverse-editing plant taxa needs to be established in the near future. As an alternative
approach, candidate editing factors could possibly be identified from the enormous genomic data that
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have been accumulated for bryophytes, lycophytes, and ferns. Subsequently, a loss-of function of those
candidates via genome editing might lead to the identification of novel editing factors.
Acknowledgments: This study was supported by JSPS KAKENHI grant numbers 2529105 and 15K14917
(to Mamoru Sugita), by JSPS Research Fellowship for Young Scientists to Mizuho Ichinose (25 3052). We thank
Sanjay Kapoor for copy editing of this manuscript.
Author Contributions: Mamoru Sugita and Mizuho Ichinose drafted, edited and wrote the paper.
Conflicts of Interest: The authors declare no conflict of interest.
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